Optimized Analysis of Proteins from Xenopus Oocytes and Embryos by Immunoblotting

Abstract

Analysis of proteins by SDS-PAGE followed by immunoblotting (western blotting) is a vital part of the molecular biologist's toolkit. This technique separates a complex protein mixture by molecular weight and then assays the presence of target proteins using specific antibodies. Immunoblotting has a variety of applications. Examples include use as a targeted approach to study protein-protein interactors or as a control to confirm expression or depletion of protein targets. However, successful execution of immunoblotting requires complicated, multistep experiments. Protocols must be optimized for each organism, target protein, and application. Therefore, knowledge gaps exist for the use of immunoblots in many models including the model frog Xenopus laevis.

Due to their large size, abundant material for biochemical experiments, and facile handling, X. laevis oocytes and embryos have been vital for studying principals of translational control. However, this species lacks specific protocols for robust and routine immunoblotting. Here, we offer an in-depth protocol for western blotting optimized for samples from multiple Xenopus developmental stages. We then analyze translational regulators across development.

SUMMARY:

This article describes a protocol for analysis of proteins from Xenopus oocytes and embryos by immunoblotting. Collection steps are described, followed by steps corresponding to sample processing, SDS-PAGE, transfer, antibody staining, and imaging. The protocol emphasizes studying translational regulatory protein complexes with endogenous antibodies and antibodies against protein affinity tags.

INTRODUCTION:

Understanding cellular mechanisms requires monitoring specific protein species. Common inquiries include how they change in expression spatially and temporally, what proteins they interact with, and if they are modified in response to different conditions. Immunoblotting, colloquially known as a “Western blotting,” is a key methodology to address these challenges. An immunoblot experiment separates proteins from a complex sample—for example, whole cell lysate—and identifies them using antibodies. Specifically, proteins are denatured and then separated by SDS-PAGE by size. The size fractionated proteins are transferred onto a solid support, such as nitrocellulose membrane, and the protein of interest is identified using antibody reagents. Immunoblotting has become a standard part of the molecular biologist’s toolkit. It is commonly used for monitoring protein expression in different contexts or as an endpoint for experiments such as immunoprecipitation assays. Despite these advantages, analysis of steady-state protein levels with immunoblotting proves challenging, as the assay must be optimized for each step and experimental context (Figure 1).

Figure 1. Workflow for Xenopus sample preparation and subsequent immunoblot analysis.

One challenging context is analyzing material from the African clawed frog Xenopus laevis. X. laevis is an important organism for studying RNA-protein interactions. This system has numerous advantages, chief among them that X. laevis produce hundreds of oocytes and subsequently synchronously developing embryos at a time. Each oocyte has ~25 ug of non-yolk protein ¹, providing abundant material for biochemical experiments. The large size (~1250 μm)¹ of oocytes and embryos also facilitates mRNA microinjection to exogenously express tagged or mutated proteins at scale². These qualities enable powerful experiments to assay translational control in X. laevis such as the tethered function assay, in which the impact of a translational regulatory protein on an mRNA can be assayed irrespective of that protein’s capacity for RNA binding ^3-6. However, immunoblotting in Xenopus oocytes and embryos comes with difficulties, including interference from large masses of yolk platelets in these early cells⁷ which will obscure results if not removed.

Here, we present an optimized protocol for effective immunoblotting in X. laevis, with an emphasis on detecting translational regulatory proteins. We provide instructions from how to collect and process oocytes and embryos to imaging the final immunoblot. Also included are detailed instructions for optimal protein loading and imaging, as well as preventing yolk contamination of the sample. In addition, we discuss possible protocol modifications and strategies to find antibodies compatible with Xenopus proteins. We intend to provide investigators guidance to effortlessly perform immunoblotting as part of their own experiments in this underutilized model organism.

PROTOCOL:

Details of equipment, reagents used, and antibody concentrations employed are listed in the Table of Materials. A selection of equipment can be viewed in Figure 2, below.

Table of Materials

Name of Material/ Equipment	Company	Catalog Number	Comments/Description
10 x Cell Lysis Buffer	Cell Signaling Technology	#9803
100 sq ft Heavy Duty Premium Wrap	Saran	2570000130
2-Mercaptoethanol	Sigma-Aldrich	M3148
2-Quart Oblong Baking/Serving Dish	Pyrex	71160010116
2x Laemmli Sample Buffer	Bio-Rad	#1610737
Anti-HA High Affinity Antibody	Roche	11867423001	Prepared as directed and used at 1:10,000 dilution.
Anti-Rabbit IgG, HRP-Linked Antibody	Cell Signaling Technology	7074S	Used at 1:30,000 dilution.
Autoradiography Cassette, 5 x 7 Inches	Research Products International	420057	Not necessary if using a digital imaging system.
Bel-Art Disposable Pestles	Millipore Sigma	BAF199230001
Bicaudal-C (Human) Rabbit Antibody	Envigo Bioproducts (now Inotiv)	N/A	Custom ordered. Used at 1:20,000 dilution.
Bicaudal-C (Xenopus) Rabbit Antibody	Envigo Bioproducts (now Inotiv)	N/A	Custom ordered. Used at 1:20,000 dilution. Ref. Park et al., 2016
Bovine Serum Albumin	Sigma-Aldrich	A3294	A substitute for nonfat milk in blocking buffer.
Centrifuge 5425	Eppendorf	13-864-456
CL-XPosure Film	ThermoFisher Scientific	34091	Not necessary if using a digital imaging system.
CNOT1 (D5M1K) Rabbit Monoclonal Antibody	Cell Signaling Technology	#44613	Used at 1:2,000 dilution.
DDX6/RCK (D26C11) Rabbit Monoclonal Antibody	Cell Signaling Technology	#8988	Used at 1:5,000 dilution.
E-Z Store & Pour Fixer	MXR Imaging	114511	Not necessary if using a digital imaging system.
E-Z Store and Pour Developer	MXR Imaging	103633	Not necessary if using a digital imaging system.
GAPDH Monoclonal Antibody	ProteinTech	60004-1-Ig	Used at 1:40,000 dilution.
Gel Releaser	Bio-Rad	#1653320
Goat anti-Mouse IgG (H+L) Secondary Antibody, HRP	Invitrogen	A16066	Prepared as directed and used at 1:40,000 dilution.
Ice Pack, 8 oz, 5 x 3 x 1"	Uline	S-18256
L-Cysteine	Sigma-Aldrich	C7352
Medical Film Processor	Konica Minolta	SRX-101A	Not necessary if using a digital imaging system.
Mini-PROTEAN Cassette Opening Lever	Bio-Rad	#4560000
Mini-PROTEAN Tetra Vertical Electrophoresis Cell	Bio-Rad	#1658004
MSE PRO 700ml Western Blot Electrophoresis Tank	MSE Supplies	BB2369
Nitrocellulose Membrane, 0.45 μm	Bio-Rad	#1620115
Nonfat Dry Milk	Sanalac	1570000180
Peroxidase AffiniPure Goat Anti-Rat IgG (H+L)	Jackson Immunoresearch	112-035-003	Prepared as directed and used at 1:100,000 dilution.
Polysorbate (Tween) 20	Fisher Bioreagents	BP337-500
Ponceau S	VWR Life Science	K793-500ML	An approximation can be made with 0.5% Ponceau S, 1% acetic acid
PowerPac Basic Power Supply	Bio-Rad	1645050
PR1MA Variable Speed 2D Rocker	MidSci	R2D-30
Precision Plus Protein Dual Color Standards Ladder	Bio-Rad	#1610374	Any protein ladder of choice can be used.
Pure Cellulose Chromatography Filter Paper, 0.35 mm	Fisher Scientific	05-714-4
Restore Western Blot Stripping Buffer	ThermoFisher Scientific	#21059
Sodium Chloride	Fisher Chemical	S640-10
Storage Box 300 mL	Rosti Mepal	RST46608	For nitrocellulose membrane blocking, antibody incubation, and washes
SuperSignal West Femto Maximum Sensitivity Substrate	ThermoFisher Scientific	#34094	An enhanced chemiluminescent (ECL) substrate containing two components: luminol and enhancer. This ECL reagent has a low-femtogram sensitivity.
SuperSignal West Pico PLUS Chemiluminescent Substrate	ThermoFisher Scientific	#34580	An enhanced chemiluminescent (ECL) substrate containing two components: luminol and enhancer. This ECL reagent has a picogram to femtogram sensitivity.
SurePAGE, Bis-Tris, 10x8, 4-12%, 10 Well Polyacrylamide Gels	GenScript	M00652
Transfer Buffer Powder	GenScript	M00652
Tris Base Ultrapure	USBiological	77-86-1
Tris-MOPS-SDS Running Buffer Powder	GenScript	M00138

Figure 2. Processed embryo extract and equipment for immunoblotting.

(A) Centrifuged X. laevis stage VI oocyte extract. Lipids and fat-soluble proteins rise to the top while vitellogenin (yolk) and pigmented debris form a pellet. (B) Equipment for SDS-page to separate proteins by molecular weight. i. Power supply, ii. Vertical electrophoresis tank, iii. Electrophoresis tank lid, iv. Electrode assembly, v. Buffer dam, vi. Precast electrophoresis gel; note the green comb (top) and blue sticker (bottom), each of which must be removed. (C) Equipment for membrane transfer. The vertical electrophoresis tank and lid are re-used for transfer after a thorough rinse. i. Roller to remove bubbles, ii. Transfer clip, iii. Small stir bar, iv. Nitrocellulose membrane in between two protective blue sheets of paper, v. Cellulose chromatography filter paper, vi. Transfer sponge cushions, vii. Glass baking dish for transfer assembly, viii. Ice pack, ix. Transfer core. (D) Other equipment. i. Gel releaser (for trimming gel wells pre-transfer), ii. Gel cassette opening lever, iii. Forceps (for handling membrane), iv. Container (for holding membrane), v. container lid.

All work with animals (X. laevis) represented in this protocol is in accordance with and has been approved by the UW-Madison Institutional Animal Care and Use Committee.

1. Collection of Xenopus oocytes and embryos

• 1.1.
  Oocyte preparation:

  • 1.1.1.
    Defolliculated X. laevis oocytes can be obtained from a commercial supplier (Ecocyte Bio Science, Austin Texas) or isolated and defolliculated as described⁸.
  • 1.1.2.
    Culture isolated oocytes in Modified Barth’s Solution (MBS ; 88 mM NaCl, 1 mM KCl, 0.4 mM CaCl₂, 0.33 mM Ca(NO₃)₂, 0.8 mM MgSO₄, 5 mM Tris-HCl, 2.4 mM NaHCO₃, pH 7.4) until use.
• 1.2.
  Embryo preparation:

  • 1.2.1.
    Isolate X. laevis eggs and fertilize them as described^{9, 10}.
  • 1.2.2.
    Culture fertilized embryos in 0.25 x Marc’s Modified Ringer’s solution¹¹ (MMR) (1 x MMR: 0.1 M NaCl, 2.0 mM KCl, 1 mM MgSO₄, 2 mM CaCl₂, 5 mM HEPES).
  • 1.2.3.
    Prepare a solution of 2% cysteine in 0.1 x MMR with the pH adjusted to 8.2 using NaOH.
  • 1.2.4.
    Remove 0.25 x MMR from the dish used for fertilization and cover the embryos with the cysteine solution to remove the jelly coat¹². Mix the solution gently so embryos are evenly exposed.
  • 1.2.5.
    Monitor embryos closely with a stereomicroscope from 2 x to 25 x magnification. After the jelly coat has dissolved (typically 3-5 minutes), the embryos will pack together closely. Avoid prolonged exposure to cysteine.
  • 1.2.6.
    Remove the cysteine solution and rinse embryos multiple times with 0.25 x MMR.
  • 1.2.7.
    Culture in 0.25 x MMR until the desired growth stage.
• 1.3.Collect the desired number of embryos or oocytes (recommended at least five per sample) in a 1.5 mL tube.
• 1.4.Remove excess media with a pipettor, avoiding damaging the cells.
• 1.5.Use samples immediately or store at −80 °C until use. Oocytes and embryos can be stored for several years.

2. Preparation of Xenopus extract

• 2.1.Thaw samples on ice if frozen.
• 2.2.Add 10 μL of chilled 10 x cell lysis buffer diluted to 1 x with double deionized water (see materials list) per embryo/oocyte and homogenize samples with a micropestle on ice.
• 2.3.Spin samples at 4 °C, 5000 x g in a benchtop centrifuge for 10 minutes to pellet debris including yolk and insoluble pigments.
• 2.4.Remove supernatant to a separate 1.5 mL tube, taking care to avoid the pellet (Figure 2A).
• 2.5.Add an equal volume of 2 x Laemmli sample buffer supplemented with 5% w/v 2-mercaptoethanol to the supernatant and boil for 10 minutes at 95-100 °C to denature sample proteins.
• 2.6.Use samples immediately or store at −20 °C for months to years.

3. SDS-Page

• 3.1.Prepare SDS running buffer: 1 x Tris-MOPS-SDS buffer (6.06 g Tris base, 10.46 g MOPS, 1.0 g SDS, 0.3 g EDTA, double deionized water to 1000 mL).
• 3.2.Remove a 4-12% bis-tris SDS precast gel from the packaging. Remove the sticker at the bottom of the gel.
• 3.3.Insert the gel into the electrode assembly opposite a buffer dam. Place the electrode assembly into the vertical electrophoresis tank (Figure 2B).
• 3.4.Fill the electrode assembly and bottom of the electrophoresis tank with 1 x Tris-MOPS-SDS running buffer.
• 3.5.Gently remove the gel comb from the gel and pipette running buffer into the wells to remove bubbles and equilibrate the buffer.
• 3.6.Carefully load 5 μL of pre-stained protein standards into the first well and 10 μL of each sample into the additional wells.

  OPTIONAL: Load remaining wells with 10 μL of 1 x Laemmli sample buffer to make the gel run more evenly.

• 3.7.Place the lid on the electrophoresis tank and connect it to a power supply. Apply 200 V.
• 3.8.Stop electrophoresis when the sample buffer dye front exits the gel.

4. Wet transfer of proteins onto membrane

• 4.1.While samples are electrophoresing, prepare 1 L of transfer buffer (3.0 g Tris Base, 4.08 g bicine, 900 mL double deionized water, 100 mL methanol). Pre-chill the transfer buffer at 4 °C.

  CAUTION: Handle concentrated methanol stocks in a fume hood and avoid contact with skin. Methanol can be substituted with ethanol.

• 4.2.
  Before the electrophoresis is complete, begin to assemble the transfer “sandwich” in the transfer clip (Figure 2C).

  • 4.2.1.
    Fill a glass baking dish with chilled transfer buffer. In subsequent sandwich assembly steps, make sure materials are submerged in this buffer.
  • 4.2.2.
    Place the transfer clip in the dish with the black half lying against the bottom of the dish, the white half open and pointed upwards, and the transfer clip open to the left.
  • 4.2.3.
    Lay one or two fiber sponges flat on the black half of the transfer clip.
  • 4.2.4.
    Cut two 0.35 mm cellulose chromatography papers (filter papers) to size and lay one on top of the sponges.
• 4.3.After electrophoresis is complete, remove the gel from the electrophoresis tank.
• 4.4.Carefully pry apart the plates of the gel with the gel cassette opening lever, ensuring that the gel remains attached to one of the plates (Figure 2D). Trim the wells from the top of the gel with the gel releaser.
• 4.5.Place the gel, still attached to one half of the plastic cassette, onto the filter paper. Remove the cassette half so the gel remains on the filter paper.
• 4.6.Cut a square of 0.45 μm nitrocellulose membrane to fit the dimensions of the gel. Remove the membrane from the protective paper, wet briefly in transfer buffer, and place it on the gel.

  NOTE: Handle the membrane carefully by the corners with gloves or forceps to avoid unwanted protein contamination of the membrane.

• 4.7.Place a second filter paper on top of the nitrocellulose membrane. Gently but firmly remove out any bubbles using a roller on top of the filter paper.
• 4.8.Stack one to two more fiber sponges on top of the filter paper.
• 4.9.Close the transfer cassette and clip it tightly shut, completing the ‘sandwich’.
• 4.10.Insert the transfer sandwich into the transfer core with the transfer clip facing upwards and the black side of the transfer clip facing the black side of the core.
• 4.11.Place the transfer core into the electrophoresis tank. Add an ice pack and a stir bar to the tank such that the stir bar can rotate freely.

  NOTE: In addition to or in place of an ice pack, the transfer can be run in a cold room at 4 °C in step 4.13.

• 4.12.Fill the tank with chilled transfer buffer. Secure lid firmly on top.
• 4.13.Connect the transfer apparatus to power supply, making sure to align electrode colors. Run the transfer for one hour, 100 V, at 4 °C, with stir bar rotating.

5. Post-transfer membrane processing

• 5.1.Prepare 10x Tris-Buffered Saline (100 mM Tris base; 1.5 M NaCl). Adjust pH to 8 and filter sterilize.
• 5.2.Prepare 1 x Tris-Buffered Saline with Tween-20 (TBSTw) by diluting 10 x TBS solution 1:10 in 500 mL double deionized water and adding 500 μL Tween-20.
• 5.3.Prepare blocking buffer (500 mL TBSTw, 25 g nonfat milk powder).

  NOTE: Nonfat milk can be substituted with 2-5% bovine serum albumin (BSA) in the blocking buffer. BSA should be used if the experimental application involves visualizing biotinylated proteins, as milk contains biotin.

• 5.4.Once transfer is complete, carefully remove and disassemble the sandwich and remove the nitrocellulose membrane. Mark which side of the membrane was contacting the gel and keep this side face up in subsequent steps.
• 5.5.Prepare 50 mL Ponceau stain (0.5% Ponceau S, 1% acetic acid), a reversible stain for protein visualization.

  CAUTION: Ponceau stain is corrosive. Avoid contact with skin.

• 5.6.Place the membrane in a small container (as shown in Figure 2D) so that it lies flush against the bottom of the container. Cover the membrane in Ponceau stain to immersion and rock gently on a rocker for 5-10 minutes.
• 5.7.Pour off the Ponceau stain.

  NOTE: Ponceau stain can be re-used multiple times.

• 5.8.Rinse the membrane in repeated washes of double deionized water until excess Ponceau is removed and protein bands in the lanes begin to resolve.
• 5.9.Confirm the efficiency of the transfer by the presence of straight, even lanes with clearly resolved bands.
• 5.10.Pour off any remaining liquid and immerse the blot in blocking buffer to prevent nonspecific antibody binding to the membrane in downstream steps.
• 5.11.Rock gently on a rocker for 30-60 minutes at room temperature.

6. Antibody incubation

• 6.1.Dilute primary antibody in 10 mL of fresh blocking buffer to the desired concentration.

  NOTE: The optimal concentration will have to be determined empirically for each new antibody. A typical starting point is 1:1000, or else follow the manufacturer’s recommendations.

• 6.2.Remove the blocking buffer and replace with the antibody mixture. Incubate, rocking gently, overnight (~16 hours) at 4 °C.
• 6.3.Pour off the primary antibody solution and rinse the membrane with TBSTw by immersing it in solution and quickly pouring it off.

  NOTE: Most antibody mixtures can be saved and reused two to three times. This must be determined empirically for each antibody.

• 6.4.Wash the membrane by immersing in TBSTw and rocking gently at room temperature for five minutes. Perform three total five minute washes.
• 6.5.Dilute the secondary antibody in 30 mL TBSTw to the desired concentration.

  NOTE: Concentration will have to be determined empirically for each antibody. We typically use 1:30,000.

• 6.6.Pour off the TBSTw wash solution and replace with the secondary antibody mixture. Incubate, rocking gently, for 45 minutes at room temperature.
• 6.7.Quickly rinse the membrane once with TBSTw followed by three five-minute TBSTw washes.

7. Imaging the membrane on a film developer

• 7.1.In a darkroom, turn on a film developer and allow it to warm up.
• 7.2.Cut two squares of plastic wrap large enough to encompass the membrane. Using forceps, place the nitrocellulose membrane ‘face up’ on the first square of plastic wrap.
• 7.3.In a 1.5 mL tube, mix equal amounts of the enhanced chemiluminescence (ECL) Luminol and Enhancer reagents. Using 1 mL of the mixed solution should cover most membranes.
• 7.4.Pipette the ECL reagent onto the membrane and manipulate the membrane gently using the plastic wrap to ensure full coverage. Incubate for one minute.
• 7.5.Remove excess ECL reagent by grasping the membrane with forceps and gently shaking off liquid or wicking away liquid by touching a corner of the membrane to a paper towel.
• 7.6.Place the membrane face up on the second square of plastic wrap and fold the plastic wrap over the membrane until it is loosely sealed. Tape the wrapped membrane securely into an autoradiography cassette.
• 7.7.In the darkroom, place a film within the cassette, covering the taped membrane. Close the cassette securely and expose the film to the membrane for one minute.
• 7.8.Carefully remove the film, being careful not to drag the exposed film along the membrane. Feed it into the film developer.
• 7.9.Upon evaluating the developed film, adjust exposure time with subsequent films until the desired protein visualization is achieved. If the protein visualization provides a weak signal, repeat the ECL reagent incubation step with a more sensitive ECL reagent and re-image.
• 7.10.Align the developed film with the glow-in-the-dark markers and use this reference to mark the position of the pre-stained protein standards on the membrane onto the film.
• 7.11.Once imagining is complete, carefully remove the membrane from the plastic wrap and immerse it in TBSTw to wash away the remaining ECL reagent.

8. (Optional) Additional antibody incubations

NOTE: Stripping an immunoblot refers to removal of the initial primary and secondary antibodies with a denaturing solution (stripping buffer) so the blot can be re-probed with a different antibody. Re-probing allows the same immunoblot to be analyzed for expression of different protein targets and to compare unknown proteins to well-characterized proteins that function as standards. Probe with the antibody that produces the comparatively weakest signal first. Doing so prevents artifacts caused by incompletely stripped antibodies in subsequent exposures and minimizes the impact of protein loss from repeated stripping of a blot.

• 8.1.Strip bound antibodies by immersing the membrane in stripping buffer and rocking gently for 5-10 minutes.
• 8.2.Remove membrane from the stripping buffer and rinse briefly in TBSTw to remove any residual stripping solution.
• 8.3.Immerse the membrane in blocking buffer and rock gently for 30 minutes.
• 8.4.Make a new solution of primary antibody in blocking buffer with the desired concentration.
• 8.5.Add the new primary antibody dilution to the membrane and incubate rocking gently overnight at 4 °C.
• 8.6.Proceed from step 6.3 onwards. The membrane can be stripped and re-probed up to three additional times.

REPRESENTATIVE RESULTS:

To demonstrate the efficiency of our immunoblotting protocol, we analyzed affinity-tagged and endogenous translational control proteins in three types of X. laevis samples: stage VI oocytes, stage 7 embryos (blastula), and stage 10.5 embryos (gastrula). These stages were chosen because they span the mid-blastula transition (stage 8.5), which marks the beginning of robust zygotic transcription ^{13, 14}. Because development prior to the mid-blastula transition occurs in absence of transcription, pre-zygotic (i.e., maternally-controlled) embryos rely heavily on translational control mechanisms to express cell fate proteins at the correct temporal and spatial resolution ¹⁵. Therefore, the mid-blastula transition is a vital context in which to study RNA-protein interactions.

To analyze protein expression via immunoblotting, X. laevis oocytes and embryos were injected with 500 pg of mRNA encoding the C-terminal half of X. laevis Bicaudal-C (Bicc1) fused to 3 x hemagglutinin (HA) affinity tags. We chose to analyze Bicc1 due to its relevance to Xenopus biology and protein-RNA interactions. Bicc1 is an mRNA-binding protein and translational repressor that guides cell fate decisions in the early embryo ^16-18. Later in development, it impacts left-right patterning ^19-21 and the function of organs including the kidneys ^22-25. Bicc1 contains a region that directs translational repression⁵ and hnRNP K homology (KH) domains that mediate binding to specific mRNAs ^{5, 26-28}.

Extracts were prepared from five HA-Bicc1-injected oocytes or embryos, along with corresponding uninjected samples as negative controls. One tenth of each extract was electrophoresed on a 4-12 % gradient bis-tris gel and wet transferred onto a nitrocellulose membrane. Ponceau staining of the membrane to detect total protein confirmed successful transfer (Figure 3A). As part of our studies, we generated an antibody in rabbits that recognizes the C-terminal half of the human Bicc1 protein. This antibody was used to detect endogenous and exogenous Xl-Bicc1 protein (Figure 3B). Previous work showed that Bicc1 protein is absent from X. laevis oocytes, but is expressed in pre-MBT embryos, before the zygotic genome is active ¹⁶. This suggests that while Bicc1 mRNA accumulates during oogenesis, it is translationally repressed. In agreement with this previous work, endogenous Bicc1 was not detected in oocytes. In contrast, the antibody recognized endogenous Bicc1 (~MW 107 kDa) in both stage 7 and 10.5 embryos. Taken together, these results validate that the antibody recognizes the endogenous Bicc1 protein (Figure 3B, top panel, red arrowhead). The Bicc1 antibody recognized a smaller protein of approximately 70 kDa in extracts prepared from mRNA-injected samples (Figure 3B, top panel, black arrowhead, compare lanes 2, 4, and 6 with lanes 1, 3, and 5). As a control for the specificity of the Bicc1 antibody, the membrane was stripped and re-probed with an antibody against the HA affinity tag. The HA antibody identified the 70 kDa protein in the injected samples but did not recognize the endogenous Bicc1 (Figure 3B, second panel, black arrowhead). Detection of the HA-Bicc1 C-term varies in intensity across stages due to differences in protein expression from injected mRNA in the different cell types.

Figure 3. Identifying levels of translational regulatory proteins through immunoblotting in X. laevis.

(A) Ponceau staining of the immunoblot to identify total protein from X. laevis stage VI oocytes, stage 7 embryos, and stage 10.5 embryos. Each lane represents 10 % of the protein prepared from an extract (0.5 oocyte or embryo). (B) Immunoblot analysis of selected X. laevis translational regulatory proteins in stage VI oocytes, stage 7 embryos, and stage 10.5 embryos. Lanes 2, 4, and 6 correspond to samples microinjected with HA-Bicc1 mRNA prior to analysis, while lanes 1, 3, and 5 serve as negative (uninjected) controls. α-Bicc1 antibody was used to assay expression of endogenous Bicc1 across stages (top panel). The blot was subsequently stripped and re-probed with an antibody to the HA affinity tag as a control for α-Bicc1 antibody specificity (second panel). Blots were stripped and re-probed for additional regulatory proteins using α-Cnot1 (third panel) and α-Ddx6 (fourth panel). α-Gapdh (bottom panel) served as a control for equal protein loading. Red arrowheads mark the location of endogenous Bicc1, while black arrowheads mark the location of exogenously expressed HA-Bicc1 C-term.

Next, we expanded our results by testing the expression of endogenous translational regulatory proteins that interact with Bicc1. First, we analyzed the expression of Ccr4-Not Transcription Complex Subunit 1 (Cnot1), a large scaffold protein and part of the Ccr4-Not deadenylase (CNOT) complex. The multi-subunit CNOT complex is one of the principal eukaryotic deadenylases and is primary known for trimming the poly(A) tail at the end of messenger mRNAs as a prelude to their degradation. However, this complex has diverse roles in translational control that extend beyond deadenylation²⁹. We were interested in assaying Cnot1 expression due the CNOT complex’s importance to mRNA regulation and previous observations that Cnot1 interacts with Bicc1^{20, 30}. To analyze Cnot1 expression, we used an antibody that recognizes the endogenous Cnot1 protein. It identified two proteins at 250 and 270 kDa in each sample, the predicted size of Cnot1 (Fig. 3B, third panel). Thus, this protocol allows us to readily identify a critical component of a regulatory complex. In addition, this data supports the protocol’s ability to successfully identify high molecular weight proteins.

To further test the generalizability of these results, we next analyzed samples for the presence of a prominent DEAD-box helicase involved in translational repression, DEAD-box helicase 6 (Ddx6, previously known as xp54 in Xenopus and me31b in Drosophila). Ddx6 is an important component of ribonucleoprotein granules, is involved in mRNA storage, and interacts with Bicc1 and Cnot1^{6, 31, 32}. An antibody recognizing the endogenous Ddx6 identified a protein of around 54 kDa, in each sample (Fig. 3B, fourth panel). Expression was reduced in zygotic embryos (Fig. 3B, compare lanes 5 and 6 to 1-4), as reported previously ³³.

Finally, to ensure that the differences we observed between samples were due to differences in expression, we stripped and re-probed the immunoblot with an antibody recognizing Glyceraldehyde 3-phosphate dehydrogenase (Gapdh) enzyme. Gapdh serves as a ubiquitously expressed “loading control.” Equivalent levels of Gapdh expression confirm the Ponceau staining: an equal amount of total protein is being analyzed across samples (Fig. 3B, fifth panel).

Together, these results confirm the efficacy of this immunoblot method. We were able to identify exogenous and endogenous Bicc1 across multiple X. laevis developmental stages relevant to the study of RNA-protein interactions: oocytes and pre-MBT embryos, which contain only maternally deposited mRNAs, and post-MBT embryos, which also include zygotically transcribed mRNAs. We were able to generalize these results by analyzing the expression of multiple translational control proteins at a variety of molecular weights (Bicc1, Cnot1, and Ddx6) and a loading control (Gapdh). We also show that this protocol can be used for X. tropicalis embryos with modification to increase the amount of material used to account for their smaller size (Supp. Fig. 1).

DISCUSSION:

Though simple in concept, immunoblotting is difficult in practice owing to multiple steps spanning several days. As such, achieving consistent, high-quality results is often challenging. Nevertheless, immunoblotting is a vital and ubiquitous component of modern biology. For example, immunoblotting is critical to confirming successful translation and expression of the microinjected mRNA in X. laevis oocytes and embryos, highlighting its importance to X. laevis research. Moreover, immunoprecipitation experiments utilize immunoblotting to analyze in vivo protein-protein interactions. Efficient detection of specific proteins will largely depend upon their abundance and the quality of the antibody detection reagents. In this study, estimated concentrations of the endogenous protein we analyzed range almost 500-fold in the Xenopus egg (Bicc1: 37.22 nM, Cnot1: 115.08 nM, Ddx6: 980.55 nM, Gapdh: 17,907.26 nM)³⁴, but all were detected readily. Below this range, modifications can be made to the workflow at essentially every step of an immunoblot—from sample collection to imaging—to improve accuracy and efficiency of detection, if needed.

Which type of SDS-PAGE gel and membrane to use are major considerations for any immunoblotting experiment. In our experience, precast gels are more facile and reproducible than gels produced in house. Precast gels have the added advantage of being available in gradient formats where the gels gradually increase in polyacrylamide concentration from top to bottom. Gradient gels allow researchers to resolve and analyze a broad range of proteins with vastly different molecular weights. Doing so avoids the typical tradeoff between choosing a low acrylamide percentage gel (<7%), which resolves high molecular weight proteins versus high percentage gel (>12%), which is better at resolving low molecular weight proteins. It is also important to consider whether to use a nitrocellulose or polyvinylidene fluoride (PVDF) membrane for protein transfer. The mechanism and advantages of each membrane type have been discussed extensively ^{35, 36}. In short, PVDF has a higher protein binding capacity (and therefore higher sensitivity) than nitrocellulose membrane and is more durable for repeated usage. However, PVDF requires additional handling in the form of activation by methanol incubation and is more expensive. While we find that both membranes are suitable for most applications, we routinely use nitrocellulose due to cost considerations.

Imaging the immunoblot is a crucial component of the protocol. During this step, researchers must exercise judgement to capture an image of an immunoblot that conveys all the necessary information but keeps background noise to a minimum. There are two main imaging methods for standard HRP-conjugated antibodies: x-ray film or digital imaging systems. In our experience, capturing images with x-ray film has many benefits. X-ray films are generally far more sensitive than digital imaging systems, facilitating the detection of low-abundance proteins. X-ray films also produce a physical record of the experiment that can be annotated and digitized at high resolution. However, a digital imaging system will have a far superior linear range–the relationship between signal strength and image intensity–than a film developer. Due to its sensitivity, an x-ray film can become overexposed quickly, losing quantitative information. Therefore, a researcher looking for precise quantitation of their results should use a digital imaging system or take special care to not overexpose developing immunoblots with x-ray film.

If immunoblotting results are unsatisfactory after following the protocol as we have outlined, such as faint or “blurry” bands upon imaging, multiple adjustments can be made to improve the outcome. For example, the amount of Xenopus protein mixture analyzed can be adjusted. Total protein from as little as one fifth of an embryo can produce strong results, though it is possible to load up to two embryos or oocytes per gel lane. Loading too much protein may result in artefacts or spurious high background signals during imaging. Additionally, electrophoresis at a lower voltage (100 V) for a longer running time generally leads to sharper protein bands. Similarly, performing the protein transfer overnight at 4°C at low voltage (30 V) instead of an hour at 100 V may improve protein detection, especially for high molecular weight species. Lastly, some primary antibodies incubations detect their targets more robustly using incubations of 1 hour at room temperature, as opposed to 4°C overnight. This protocol modification must be tested for each primary antibody.

Identifying antibodies to detect endogenous proteins presents a major challenge to immunoblotting in Xenopus, as relatively few Xenopus-specific antibodies are commercially available. Researchers can benefit from non-commercial sources of Xenopus antibodies such as Xenbase³⁷ (https://www.xenbase.org/xenbase/), the Developmental Studies Hybridoma Bank (https://dshb.biology.uiowa.edu/), and the European Xenopus Resource Centre (https://xenopusresource.org/). Alternatively, antibodies generated to recognize human or mouse protein may be useful for detecting homologous Xenopus proteins, depending upon the conservation of the regions used for antibody production. For example, a Bicc1 antibody presented in this protocol was raised against human Bicc1 protein. For such reagents, the ability to cross-react with Xenopus proteins must be determined empirically. Though the exact epitope companies use to create antibodies are generally proprietary, many will provide an amino acid range for the epitope upon request. This region can then be aligned with the corresponding region of the homologous Xenopus protein. While success is not guaranteed, we have typically had success when the epitope used for antibody production in the target species exhibits at least 75 % or above similarity to the comparable stretch of sequence in Xenopus. Antibodies generated against non-Xenopus proteins should be validated before extensive use, as improper verification of antibodies is a major contributor to irreproducible research^{38, 39}. This can be done by expressing a tagged Xenopus protein of interest in embryos through mRNA microinjection⁴⁰. Analyzing the same samples with an antibody towards the affinity tag is then used to confirm that the antibody to the endogenous protein and the antibody to the tagged protein both recognize the same entity (as in Figure 3B).

Analysis of RNA-binding proteins, especially those with regulatory functions such as Bicc1, can prove challenging due to a dearth of reagents and typically low expression of potent regulatory molecules. This problem becomes more apparent in immunoblotting, where each reagent must be optimized separately, and finding antibodies of sufficient specificity and sensitivity may be difficult. Our intent with this protocol is to address specific limitations of immunoblotting in Xenopus and elaborate on steps applicable to other systems such that researchers may overcome the difficulties of analyzing RNA-binding proteins.

Supplementary Material

Sup Fig.

Supplemental Figure 1. Immunoblot analysis of Bicc1 protein in Xenopus laevis versus Xenopus tropicalis. α-Bicc1 antibody was used to assay expression of endogenous and exogenous Bicc1 across different amounts of X. laevis and X. tropicalis embryo extract. Lane 1: 0.5 uninjected X. laevis embryo. Lane 2: 0.5 HA-Bicc1 injected X. laevis embryo. Lane 3: 3 uninjected X. tropicalis embryos. Lane 4: 1 uninjected X. tropicalis embryo. Lane 5: 0.5 uninjected X. tropicalis embryo. Two antibodies recognizing endogenous Bicc1 were used: one raised against human Bicc1 (top panel) and the other raised against X. laevis Bicc1 (middle panel). α-Gapdh (bottom panel) was used to control for equal protein loading.